Phytate is the major storage compound of phosphate in plant seeds and binds up to 80% of the total phosphate content in cereal grains (Eeckhout and Depaepe, 1994). It consists of a six-carbon ring with six phosphate groups attached. The negative charge is balanced by cations of magnesium, calcium among others. Together they form large crystals that are stored within the aleurone embryo or the endosperm of the seed.
Monogastric animals generally are not able to degrade the phytate present in the feed and the phosphate compound is thus, through the manure, administered to the environment. This leads to eutrophication of lakes, streams and the coastal sea, which results in increased growth of algae and in the end to sub-oxygen regimes and the death of aquatic life. For people with unbalanced diets as seen in many undeveloped countries the insufficient digestion of phytate is a severe nutritional problem, particularly because it is sequestering zinc and iron from uptake. The formation of insoluble aggregates of phytate with important minerals as zinc and iron as well as with proteins leads to poor digestibility of all the agents.
The scientific interest in phytate and its metabolic enzymes goes back more than a century, though it probably received the most attention from the general public in the early seventies. With the reintroduction of vegetable diets and wholemeal bread the degradation of phytate became a problem for human nutrition in the western world. Today phytate related concerns in the western world involves the negative effects on the environment caused by the intensive production of fish, pigs and poultry.
Phytate is the trivial name for the mixed salt of 1,2,3,4,5,6 myo-inositol-hexakisphosphate or phytic acid (InsP6). Myo-inositol is synthesised from D-glucose via three enzymatic steps, a) hexokinase (EC 2.7.1.1), b) 1L-myo-inositol 1-phosphate synthase (EC 5.5.1.4) and c) myo-inositol 1-phosphate phosphatase (Loewus and Murthy, 2000).
Phytate (InsP6) is believed to function as an effective storage compound in the seed of both phosphate and essential cations, especially potassium and magnesium. The inositol moiety, the phosphate groups as well as the chelated cations are believed to be utilised by the growing seedling.
Where (InsP6 in the plant cells have been assigned a pure storage function and as a precursor of the lower InsP, recent reports have shown InsP6 to act as a signalling molecule (Voglmaier et al., 1992), (Larsson et al., 1997) in animal systems, in yeast (York et al., 1999) and in plants (Munnik et al., 1998), (Muir and Sanders, 1997)
Phytases are a group of phosphatases that catalysis the stepwise removal of ortophosphate from phytate. Phytase enzymes are classified into two groups according to the initial position of hydrolysis. All the fungal phytases investigated as well as the novel phytases from Bacillus subtilis and B. amyloliquefaciens (Kerovuo et al., 1998 & 2000; Kim et al., 1998) initiate the hydrolysis of phytate at position 3 (EC 3.1.3.8) and catalyse the reaction:

(The Bacillus enzyme is stated in the EMBL accession as 3-phytase but it has not been published elsewhere). The plant phytases as well as the enzyme from E. coli are 6-phytases (EC 3.1.3.26) and catalyse the reaction:

In this instance the L-configuration is used and the removed phospho-group in the latter reaction scheme is situated at position 4 when the D-configuration is assigned.
Most phytases identified are enzymes that accept a broad range of substrates and as such, phytases are a rather loosely defined subclass of phosphatases.
Numerous phytase enzymes hav been characterised from fungal, bacterial, animal and plant sources (Dvorakova, 1998). However, microbiel phytase enzymes are by far the best known. These include Aspergillus niger PhyB (Ehrlich et al., 1993), A. fumigatus (Ullah and Dischinger, 1993), A. niger PhyA (van Hartingsveldt et al., 1993), A. niger w. awamori (Piddington et al., 1993), A. terreus (Mitchell et al., 1997), A. ficcum (Ullah and Dischinger, 1993), Emericella nidulans (Aspergillus nidulans) (Mitchell et al., 1997), the heat tolerant Talaromyces thermophilus (Pasamontes et al., 1997), E. coli (Jia et al., 1998), Bacillus sp. (Kim et al., 1998) and Bacillus subtilis (Kerovuo et al., 1998).
Among plants phytase activity from wheat, rye, spelt, oat, rice and maize have all been subjected to purification and characterisation procedures. Phytases have been characterised from different plant tissues but only the exceptional alkaline phytase from lily pollen has phytate as the sole substrate (Scott and Loewus, 1986; Baldi et al., 1988; Barrientos et al., 1994).
In 1997 the first plant phytase was cloned from Zea maize (Maugenest et al., 1997). This enzyme was initially purified in 1993 (Laboure et al., 1993) and was described as a homo-dimer of a 38 kD polypeptide. An expression library was screened with antibodies raised against the purified protein and this lead to the identification of a full-length cDNA clone (phyS11). Two peptide sequences determined from the purified maize enzyme were encoded by the cDNA clone. The phyS11 clone was expressed in E. coli and a polypeptide with the same migration, when using SDS-PAGE and native-PAGE, as the maize phytase was obtained. However, no phytase activity associated with the heterologous polypeptide could be detected, even when the E. coli expressed protein was applied in 10 times higher concentrations than the detection level of the native enzyme activity (Maugenest et al., 1997). Neither has genetic transformation of maize with phyS11 constructs resulted in expression of an active phytase enzyme (P. Perez Limagrain, pers. Comm.).
The soybean phytase purified to apparent homogeneity from 10-day-old germinating cotelydons (Gibson and Ullah, 1988) is a monomeric enzyme with a native molecular w ight of 50 kD and it migrat s as two bands of 59 and 60 kD during SDS-PAGE (see tabl 1.). The enzyme activity is strongly inhibited by phosphate and 0.5 μM phosphate renders the enzyme only 67% active (apparent Ki=18 μM). This implies that the enzyme activity is tightly regulated by product inhibition.
The soybean phytase is reportedly blocked N-terminally, but an internal 18 amino acid from the enzyme was published in a 1990 review (Gibson and Ullah, 1990). This sequence (MHADQDYCANPQKYNXAI (SEQ ID NO: 13)) matches 100% with the sequence of soybean β-amylase (result not shown).
Another, N-terminal, soybean phytase sequence was published in GB 2319030. The sequence disclosed in GB 2319030 is similar to enzymes of the purple acid phosphatases (PAP) family, however this is not described in the patent.
Phytase enzymes from wheat bran having an activity optimum at pH 5.0 were purified and studied in detail by Nagai and Funahashi in the early sixties (Nagai and Funahashi, 1962; Nagai and Funahashi, 1963). Ten years later Lim and Tate (Lim and Tate, 1971; Lim and Tate, 1973) further published the presence and partial purification of two wheat bran phytase enzymes that could be separated on DEAE-cellulose but with identical molecular weights of 47 kD. The two enzyme fractions F1 and F2 differed by their pH optimum for activity of 5.6 and 7.2, respectively. The phytase activity of fraction F1 with a pH optimum at 5.6 (Lim and Tate, 1973) had many similarities to the earlier described phytase activity (Nagai and Funahashi, 1963) with an optimum at about 5.0, but the F1 activity was inhibited by phosphate and the latter was not inhibited at all. The products from the hydrolysis of phytate by the F1 and F2 enzyme preparations were analysed (Lim and Tate, 1973) and it was found that although 6-phytase (the product being D-ins1,2,3,5,6) was the primary activity of both fractions, the F2 fraction also exhibited 5- and 2-phytase activity (Irving, 1980).
Nakano et al., 1999 purified and biochemically characterized two wheat phytase enzymes from the “Nourin 61” wheat variety. The N-terminal sequence of both enzymes were determined to be 13 amino acid residues long having one unknown amino acid. A Swiss-Plot database examination did not reveal homologue sequences. The inv ntors did not disclose any gene or cDNA sequences.
In 1997 Nakano et al. purifi d and biochemically characterized three N-terminal sequences from wheat bran isoenzymes. Homologue sequences, gene or cDNA sequences were not described.
Any PCR based cloning strategy requires both a forward and a reverse primer. The N-terminal amino acid sequence can potentially be used to construct a forward primer, but no specific reverse primer can be made. It is therefore necessary to use a strategy using one unspecific primer in combination with the specific forward primer. Although such strategies exist (WALK-PCR, TAIL-PCR) they do far from always prove successful and more importantly they have as an absolute requirement that at least two nested non-degenerate primers can be constructed from the known sequence, which is not possible from the N-terminal amino acid sequence published by Nakano et al. 1999. This is because of the high codon degeneracy of the amino acids present in the amino terminal polypeptide (N-terminal sequence of Nakano et al.). For example, the amino acids arginine, serine and leucine are each represented by six codons in the standard genetic code, while the amino acids valine, threonine, proline, glycine, alanine are represented by four codons each. Furthermore, the fourth residue from the N-terminal is unknown Xaa, which altogether means these amino acids constitute 12 of the 13 residues in the amino terminal polypeptide sequence. Reverse transcription coupled PCR, RT-PCR on mRNA is inefficient and by no means trivial for fragment sizes over 700 bases, thus demanding internal sequences for primer design.
Alternative strategies are based on hybridisation screening of cDNA or genomic libraries, but a highly degenerate oligonucleotide of a maximum of 39 bases is not sufficient to do this
Although it has been the aim of several researchers to obtain phytase sequence information from cereals, such as wheat and barley, it is not until now that an isolated phytase from wheat is presented, and specific wheat phytase sequences are disclosed along with cloned wheat and barley phytase encoding nucleotide sequences.